Protocol for optical clearing and imaging of fluorescently labeled ex vivo rat brain slices

Summary Tissue clearing is commonly used for whole-brain imaging but seldom used for brain slices. Here, we present a simple protocol to slice, immunostain, and clear sections of adult rat brains for subsequent high-resolution confocal imaging. The protocol does not require toxic reagents or specialized equipment. We also provide instructions for culturing of rat brain slices free floating on permeable culture inserts, maintained in regular CO2 incubators, and handled only at media change.


4-(2-Hydroxyethyl)-1-piperazineethanesulfonic acid, HEPES
CRITICAL: The solution must be prepared on the day of the experiment and saturated with carbogen (95% O 2 / 5% CO 2 ) for at least 10 min and stored at 4 C.
CRITICAL: The solution must be freshly prepared, sterilized by filtration through a 0.2 mm syringe filter unit and equilibrated to 5% CO 2 at 37 C for at least 30 min before use. This solution is suitable for storage at 4 C and can used within two weeks if L-Glutamine 200 mM is freshly added every time before use.
Alternatives: L-Glutamine 200 mM can be substituted with 1% GlutaMAX Supplement (100X) which is more stable and does not spontaneously degrade. This solution can be stored ad 4 C and used for one week.
Note: This table describes the volume required for storing 10 brain sections, considering 1 mL per section.
Note: This table describes the volume required for 10 brain sections, considering 1 mL per section at each incubation step (permeabilization, primary antibody, secondary antibody conjugated to fluorescent-dyes).

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Alternatives: Use serum specific to the host animal that the secondary antibody was generated in (i.e., goat serum for goat anti-rabbit secondary antibodies). Note that the serum cannot be from the same host species of the primary antibody (i.e., goat serum cannot be used for goat primary antibodies).
CRITICAL: Methanol is hazardous (see safety information and chemical handling) and needs to be always handled under a fume hood and with protective measures.
CRITICAL: DCM is hazardous (see safety information and chemical handling) and needs to be always handled under a fume hood and with protective measures.
Note: This table describes the volume required for 10 brain sections, considering 1 mL per section. STEP-BY-STEP METHOD DETAILS Acute brain slices preparation

Timing: 1 h
As this protocol is designed to prepare acute brain slices for culturing, it is important to autoclave all surgical instruments and materials before use. Wearing protective gloves, safety goggles, protective clothing, and a face mask is required.
1. Setting up the cutting station. a. To reduce contamination, use water and then 70% ethanol to clean all surfaces including benchtop, vibratome, buffer tray, and tubing system used for diffusing the carbogen. Submerge the razor blade in 70% ethanol and let it dry for 5 min. b. Vibratome set up.
i. Insert the razor blade into the blade holder and turn the vibratome on. ii. Check the blade's position using the VibroCheck (see Leica VT1200 S user manual, pages 34-39). iii. Install the double-walled buffer tray (Leica VT1200 S user manual, pages 34-35) and use a silicone tube to connect a gas diffuser to the cylinder of compressed carbogen. The gas diffuser can be either a small micro-filter-candle or an aquarium air curtain. iv. Turn on the cooling unit connected to the buffer tray and set it up to 3 C at least 10 min before starting the preparation. c. Fill the buffer tray with 300 mL of pre-chilled NMDG-HEPES aCSF solution and maintain it with low carbogen bubbling to maintain minimal turbulence inside the buffer tray during the cutting. d. Place the vibratome's specimen plate, a straight forceps, a Moria perforated spoon, a glass petri dish, a single Whatman round filter paper, the liquid super glue, and one razor blade on the benchtop next to the vibratome ( Figure 1A). 2. Brain dissection and vibratome sectioning.
a. Before starting the brain dissection. i. Prepare straight sharp-blunt scissors, Mayo-Noble scissors, straight bone cutter and longround tip spatula ( Figure 1B). ii. Add 80 mL of pre-chilled NMDG-HEPES aCSF solution into a 100 mL beaker, keep it inside a bucket with ice and continue carbogen bubbling using a small micro-filter-candle. iii. Prepare 50 mL of wash buffer and keep it in continuous carbogen bubbling using a microfilter-candle. b. Surgery.
i. Decapitation of the animal is performed with a rodent guillotine following depth anesthesia using an intraperitoneal injection with pentobarbital sodium (30-50 mg/kg). ii. Take the animal's head and remove the skin of the scalp using straight sharp-blunt scissors. iii. Using straight sharp-blunt scissors, make a 1 cm lateral/ventral incision cutting both sides of basioccipital bone through the foramen magnum at the occipital bone. iv. Insert the scissors again into the foramen magnum and cut dorsally alone the middle line of the interparietal, parietal, and frontal bones until it reaches the nasal bone. The cutting must be superficial and carefully performed to avoid damaging the underlying brain. Take Mayo-Noble scissors and cut the nasal bone transversally at the level of the premaxilla. v. Use the straight bone cutter to remove the nasal, frontal, parietal, and interparietal bones on both sides, starting from the rostral towards the caudal direction, to expose the brain. vi. Rinse the brain with 10 mL of pre-chilled NMDG-HEPES aCSF saturated with carbogen and then take a long-round tip spatula to gently scoop out the intact brain into the beaker prepared in step 2a. c. Brain slicing.
i. Take the brain with a Moria perforated spoon and place it on a Whatman round filter paper in a 100 3 15 mm glass petri dish to prepare a tissue block. The tissue block can be prepared by cutting the brains at different angles depending on the region of interest. To prepare cortical coronal slices, the olfactory bulb and the cerebellum were removed, and the brain glued up vertically. ii. Lift the brain tissue block with straight forceps and fix it on the specimen plate with enough liquid super glue. Immediately, transfer the specimen plate into the buffer tray filled with chilled NMDG-HEPES aCSF continuously saturated with carbogen. iii. Use the vibratome's control panel to set up the desired speed, amplitude of the vibration, and thickness of the slices which can be from 250 to 300 mm. The following parameters were used in this protocol: 0.14 mm/s speed, 1.10 mm amplitude and 275 mm slice thickness. The sectioning started from ventral towards dorsal part of the brain ( Figure 1C).
CRITICAL: all animal procedures must follow the policies and guidelines established in your organization (university or company). CRITICAL: Wear safety goggles to avoid accidents with the bone shards during the brain dissection.
CRITICAL: The tissue block preparation cannot take more than 30 sec; otherwise, it will affect the quality of the slices.

Brain slice culturing
Timing: 30-45 min After brain slicing, transfer the brain slices from the vibratome's buffer tray to a 100 3 15 mm petri dish containing oxygenated pre-chilled wash buffer using a paintbrush n 6 and take them into a laminar flow hood.
3. Preparation of equipment and buffers for brain slice culture. a. Ensure that the following necessary equipment is organized for quick access: 6-well culture plate, Millicell culture inserts, autoclaved cover glass forceps and a paintbrush n 6 for transferring the brain slices. b. Clean the brush with 70% ethanol, let it dry for 5 min and rinse with deionized water thoroughly. c. Add 1 mL of brain slice culture medium to each well of a 6-well culture plate. d. Place a Millicell culture insert into each well using flat cover glass forceps and add an additional 0.5 mL of slice culture media on top of each insert. e. Place the 6-well culture plate in the incubator at 37 C with 5% CO 2 for at least 30 min to ensure that the culture medium is pre-warmed before brain slices are plated. 4. Brain slice culture preparation.
(C) Vibratome sectioning. Silicone tube (1) is connected to an aquarium air curtain (2) surrounding the buffer tray (3). Connection of the cooling unit to the buffer tray (4). Vibratome's specimen plate submerged into the chilled NMDG-HEPES aCSF solution holding the brain tissue block (5).

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b. Manually transfer individual brain slices to each culture insert in 6-well culture plate using the paintbrush n 6. Perform this step carefully to avoid damaging the tissue (Figure 2, Methods video S1). c. Return the 6-well culture plate now containing the brain slices in culture medium to the 37 C incubator with 5% CO 2 .
CRITICAL: Make sure that no areas of the brain slices are folded or wrinkled.
Note: Brain slices submerged with media do not attach to the membrane of the Millicell culture insert and remain free-floating.
5. Brain slice culture maintenance. a. Every second day, remove 1 mL of culture medium from the bottom of each well and the excess medium that collects in the Millicell culture insert without disturbing the brain slice. b. Add 1 mL of fresh and pre-warmed culture medium to the bottom of each well and an additional 0.3 mL inside the Millicell culture insert to submerge the brain slice. c. Maintain brain slices in culture as long as required by the experimental plan.
CRITICAL: The culture medium needs to equilibrate at 37 C with 5% CO 2 for 30 min before transferring the brain slices and before every medium change. Maintain the brain slices in culture under sterile conditions. Note: (1) Transfer an entire rat coronal brain slice per Millicell culture insert or 1-2 coronal brain slice-separated hemispheres depending on your unique experimental conditions. If more than 1 brain slice is plated, avoid slice overlap and ensure that slices do not contact the sides of the insert. (2) In our experience, brain slices from 275 mm thickness are not disturbed by using a paintbrush to transfer them. Alternatively, an inverted glass Pasteur pipette can be used to position individual slices in each Millicell culture insert although this will transfer wash buffer to the culture plate. Excess wash buffer can be removed by replacing the culture media with fresh and pre-warned culture media immediately after transferring. It is important to avoid prolonged exposure to the wash buffer in culture since it can affect brain slice health. Timing: 3-4 days (for step 8) 6. Fixation of brain slices.

Immunostaining of brain slice cultures
a. Remove the culture medium and add 1 mL of 4% paraformaldehyde (PFA) solution pH 7.4 into the well and an additional 1 mL on top of the brain slice and fix for 20 min at room temperature (20 C-23 C). b. Gently transfer each fixed brain slice from the Millicell culture insert to a glass vial with lid containing 1 mL KPBS using a paintbrush. This will ensure compatibility with clearing reagents that will be used in the next steps. c. Wash the slice 2 3 5 min with KPBS at room temperature (20 C-23 C) by adding 1 mL of KPBS per vial. Note: Ensure that the volume of fixative is sufficient to fully immerse the brain slice.
CRITICAL: Do not fix brain slices longer than recommended. Immunostaining can otherwise be compromised due to excessive cross-linking of proteins. PFA is hazardous (see safety information and chemical handling) and needs to be always handled under a fume hood and with protective measures.
Pause Point: The fixed and washed brain slices can be stored at 4 C in bacteriostatic preservative (0.01% sodium azide in KPBS). Samples are stable for a few weeks. 7. Permeabilization and blocking of brain slices.
a. Remove the KPBS. b. Permeabilize and block the brain slice by adding 1 mL of blocking solution in each vial. c. Incubate overnight at room temperature (20 C-23 C), or at 4 C for two days. 8. Antibody staining of brain slice.
a. Prepare the primary antibody solution diluted (for this experiment rabbit anti-Iba1, 1:1000) in 1 mL of blocking solution. b. Remove the blocking solution from the glass vial containing the brain slice.  Pause Point: The immunostained and washed brain slices can be stored at 4 C in bacteriostatic preservative (0.01% sodium azide in KPBS) while protected from light for up to two weeks. However, we recommend continuing with clearing and imaging right after staining because the fluorescence decreases over time.
CRITICAL: Throughout the entire staining procedure, be careful not to disrupt the brain slices when removing and adding solutions. Avoid drying out the brain slices. Perform each step on a mildly shaking platform (100 RPM) for homogeneous permeabilization and staining of the tissue. Seal the glass vial during incubation using a lid or parafilm to avoid excess evaporation.

Timing: 1 day
After staining, the sample is ready to be cleared for imaging of the entire brain slice. The steps in this process are intended to remove water and lipids from the tissue and match the refractive index throughout the tissue. 9. Dehydrate brain slices in methanol.
a. Prepare the methanol series (20%, 40%, 60%, 80%, 100%). b. Remove the KPBS from the glass vial containing the brain slice. c. Add 1 mL of 20% methanol and incubate 10 min at room temperature (20 C-23 C) on a shaking platform at mild speed (100 RPM). d. Remove the methanol solution and add 1 mL 40% methanol. Incubate for 10 min at room temperature (20 C-23 C) on a shaking platform at mild speed (100 RPM). e. Remove the methanol solution and add 1 mL 60% methanol. Incubate for 10 min at room temperature (20 C-23 C) on a shaking platform at mild speed (100 RPM). f. Remove the methanol solution and add 1 mL 80% methanol. Incubate for 10 min at room temperature (20 C-23 C) on a shaking platform at mild speed (100 RPM). g. Remove the methanol solution and add 1 mL 100% methanol. Incubate for 10 min at room temperature (20 C-23 C) on a shaking platform at mild speed (100 RPM).
CRITICAL: Methanol is hazardous (see safety information and chemical handling) and needs to be always handled under a fume hood and with protective measures. Close the glass vials with the lid before positioning them on the shaking platform outside the fume hood.  CRITICAL: DCM is hazardous (see safety information and chemical handling) and needs to be always handled under a fume hood and with protective measures. Close the glass vials with the lid before positioning them on the shaking platform outside the fume hood.
11. Clear the tissue using ethyl cinnamate. a. Add 500 mL of ethyl cinnamate per well in the black Ibidi m-Plate 24 well plate with flat and clear bottom for high throughput microscopy. b. Remove the DCM from the glass vial containing the brain slice. c. Add 500 mL of ethyl cinnamate to avoid DCM carry-over. d. Gently transfer the cleared brain slice from the glass vial to the Ibidi m-Plate 24 well plate containing ethyl cinnamate using a paintbrush. e. Incubate overnight at 4 C protected from light.
Note: The tissue becomes transparent after adding ethyl cinnamate ( Figures 3A and 3B). Pause Point: The cleared brain slices can be stored at 4 C in ethyl cinnamate for weeks. However, we recommend imaging the samples the day after clearing because the fluorescence decreases over time.
CRITICAL: Be very careful not to dry the sample when changing solutions or transferring the tissue.

Confocal imaging of cleared brain slices
Timing: 5-30 min per slice We used Leica TCS SP8 point laser scanning confocal microscope with a 10x objective, but the prepared samples can be imaged with any inverted laser scanning confocal microscope equipped with appropriate objectives and filter sets and the imaging parameters would vary depending on the performed immunostaining and experimental goals. Here we used anti-Iba1 primary antibody in combination with Alexa Fluor 647 secondary antibody to visualize microglia in striatal slices. Leica dynamic filter was used for imaging. Imaging the sample using air objective or immersion objective with RI different from ethyl cinnamate (RI = 1.56) results in refractive index mismatch. Geometric distortion along the z-axis caused by RI mismatch can be correct via the formula provided below.
12. Flatten the immunostained and cleared brain slice.
a. Reduce the volume of ethyl cinnamate so it just covers the brain slice. b. Place a round glass coverslip in the well on top of the brain slice. c. Place 3-4 metal nuts on top of the coverslip amounting to 7-10 g of weight in total to mechanically flatten the brain slice. 13. Image the brain slice by taking a z-stack throughout the thickness of the sample. CRITICAL: Metal nuts might reflect light and therefore impede imaging if a spinning disk confocal or a widefield microscopes are used. In this case, remove the nuts before imaging. In our experience, brain slices remain flattened if imaged right after the removal of the weight.
Note: Care should be taken in handling the plates as the sample could move in the wells. We have not experienced brain slices moving during imaging.

EXPECTED OUTCOMES
The first part of the protocol provides detailed instructions on how to prepare 275 mm thick adult rat brain slices ( Figure 1) and place them in organotypic culture ( Figure 2, Methods video S1). Proceeding with organotypic culture is optional as brain slices can be used for electrophysiological measurements immediately after cutting and fixed once the measurements are executed. The second part of the protocol provides detailed instructions and troubleshooting guidance for fluorescence immunostaining, clearing (Figure 3), flattening and confocal imaging of fixed brain slices (Figure 4). Importantly, the protocol takes advantage of widely available reagents and tools. Furthermore, ethyl cinnamate is used for refractive index-matching which is non-toxic and therefore allows imaging on multi-user microscopes without the need for an exhaust system. Therefore, we believe that the protocol can be easily adapted by other research groups and readily integrated into experimental plans.

LIMITATIONS
Sample dehydration using methanol significantly reduces the intensity of endogenous fluorescence in 488 nm band. If retention of endogenous fluorescence is of crucial importance, 1-propanol adjusted to alkaline pH should be used for dehydration instead. 9 Quantitative morphometric analysis of cleared brain slices might be affected by distortions caused by dehydrated tissue shrinkage. Furthermore, refractive index mismatch caused by using air objectives to image samples in ethyl cinnamate leads to geometrical distortion of the final image. Due to this effect, the sample will appear shortened along the z-axis and needs to be corrected for. TROUBLESHOOTING Problem 1 Low brain slice quality is a very common problem in electrophysiology and ex vivo cultures (associated with sections: ''Acute brain slice preparation'' steps n 1-2 and ''Brain slice culturing'' steps n [3][4][5]. It can be due to several factors including osmolarity and pH of the buffer solution, oxygen deprivation, mechanical stress on the slices, and technical skills.

Potential solution
Osmolarity and pH: be sure that all devices are properly calibrated before use.
Oxygen deprivation: this occurs during brain dissection and tissue block preparation; therefore, those steps should be carried out as quick as possible. It can be resolved by improving technical skills. . Flattening of cleared brain slices (A) 3D reconstruction of an image stack displaying a section of a cleared brain slice before (left) and after flattening (right). Flattening is an important step because distortions in brain slice geometry that occur during cutting and clearing (due to variable shrinkage rates of different brain tissue areas) lead to difficulties and limitations during image acquisition. (B) Images show an Iba1-immunostained microglial cells within the brain slice before (left) and after (right) clearing procedure was performed. Orthogonal view demonstrates how much information is lost in non-cleared tissue. Clearing allows imaging of the entire thickness of the brain slice. Methods video S1: Video showing how to use a paintbrush to transfer brain slices from petri dish to 6-well culture plate containing Millicell culture inserts. Related to ''Brain slice culturing'' step 4b.

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Mechanical stress: lift the slices carefully with the paintbrush and avoid disturbing the slices excessively while performing media changes.

Problem 2
Strong staining only at tissue edges or antibody does not penetrate deeply into the tissue (associated with section: ''Immunostaining of brain slice cultures'' steps n [6][7][8]. This problem may be attributed to the permeabilization step or antibody incubation time.

Problem 3
High background signal and/or bright dots of staining in the tissue (associated with section: ''Immunostaining of brain slice cultures'' steps n [6][7][8]. This problem may be attributed to the precipitates present in PFA solution, insufficient blocking of the tissue, inadequate washing or improper (excessive) antibody concentration and/or incubation. The problem could also be contributed to residual presence of erythrocytes in microcirculation that can produce autofluorescence. If this is the case, it is recommended to perform saline perfusion during preparation of acute brain slices in order to clear the blood from the brain.

Potential solution
It is recommended to filter the PFA solution through a 0.45 mm syringe filter unit to remove any particulate matter before fixation of the brain slices. Extend the time of blocking solution incubation and/or increase the frequency of washing steps after each antibody incubation step. Titer the antibody concentration upon first use. Centrifuge the secondary antibody stock.

Problem 4
Extremely bright staining of the tissue sample (associated with section: ''Clearing of immunolabelled brain slices'' steps n 9-11). This problem may be attributed to excessive drying/shrinkage of the sample when changing solutions or transferring the tissue during clearing.

Potential solution
Leave a small amount of liquid left in the vial and apply the new solution immediately after removal.

Problem 5
Tissue samples become opaque during long term storage (associated with section ''Clearing of immunolabelled brain slices'' steps n 9-11). This issue might occur if there are aqueous solution (e.g., KPBS) in other wells in the same plate as the cleared tissue or the stored sample is exposed to increased humidity.

Potential solution
Make sure that the plate with cleared samples only contains ethyl cinnamate. Seal the plate with parafilm.

RESOURCE AVAILABILITY
Lead contact Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Janko Kajtez (janko.kajtez@med.lu.se).

Materials availability
This study did not generate new unique materials. All materials are purchasable. Data and code availability All data needed to evaluate the conclusions in the paper are present in the paper. Additional data and codes related to this paper may be requested from the authors.